The increased reliance on glutamine catabolism by proliferating cancer cells, termed glutamine addiction, has recently attracted significant attention as a route to developing new therapeutics that target this unique metabolic requirement of transformed cells (Hensley et al., “Glutamine and Cancer: Cell Biology, Physiology, and Clinical Opportunities,” J. Clin. Invest. 123:3678-84 (2013); DeBerardinis & Cheng, “Q's Next: The Diverse Functions of Glutamine in Metabolism, Cell Biology and Cancer,” Oncogene 29:313-24 (2010); Krall & Christofk, “Rethinking Glutamine Addiction,” Nat. Cell Biol. 17:1515-17 (2015); DeBerardinis et al., “The Biology of Cancer: Metabolic Reprogramming Fuels Cell Growth and Proliferation,” Cell Metab. 7:11-20 (2008)). The outcome of elevated glutamine metabolism leads to increases in glutamine-fueled anaplerosis, where glutamine is first deamidated by the mitochondrial enzyme glutaminase to produce stoichiometric amounts of glutamate and ammonia. Glutamate is then deamidated by glutamate dehydrogenase (GDH) or by one of two transaminases (GOT/GPT), producing α-ketoglutarate, which is incorporated into the TCA cycle. In this way, glutamine, being the most abundant amino acid in the blood, acts as a primary carbon and nitrogen source for highly proliferative cells. Increased glutamine metabolism is triggered by several signal transduction pathways, including those influenced by HIF1α, Myc, and Rho GTPases, as well as by Ras/MAPK- and mechanistic target of rapamycin (mTOR)/Akt-signaling activities (Sun & Denko, “Hypoxic Regulation of Glutamine Metabolism Through HIF1 and SIAH2 Supports Lipid Synthesis That Is Necessary for Tumor Growth,” Cell Metab. 19:285-92 (2014); Wise et al., “Hypoxia Promotes Isocitrate Dehydrogenase-Dependent Carboxylation of α-Ketoglutarate to Citrate to Support Cell Growth and Viability,” Proc. Natl. Acad. Sci. U.S.A. 108:19611-16 (2011); Gao et al., “c-Myc Suppression of miR-23a/b Enhances Mitochondrial Glutaminase Expression and Glutamine Metabolism,” Nature 458:762-65 (2009); Wise et al., “Myc Regulates a Transcriptional Program That Stimulates Mitochondrial Glutaminolysis and Leads to Glutamine Addiction,” Proc. Natl. Acad. Sci. U.S.A. 105:18782-87 (2008); Wang et al., “Targeting Mitochondrial Glutaminase Activity Inhibits Oncogenic Transformation,” Cancer Cell 18:207-19 (2010); Stalnecker et al., “Mechanism by Which a Recently Discovered Allosteric Inhibitor Blocks Glutamine Metabolism in Transformed Cells,” Proc. Natl. Acad. Sci. U.S.A. 112:394-99 (2015); Thangavelu et al., “Structural Basis for the Allosteric Inhibitory Mechanism of Human Kidney-Type Glutaminase (KGA) and Its Regulation by Raf-Mek-Erk Signaling in Cancer Cell Metabolism,” Proc. Natl. Acad. Sci. U.S.A. 109:7705-10 (2012); Duran et al., “Glutaminolysis Activates Rag-mTORC1 Signaling,” Mol. Cell 47:349-58 (2012); Csibi et al., “The mTORC1/S6K1 Pathway Regulates Glutamine Metabolism Through the eIF4B-Dependent Control of c-Myc Translation,” Curr. Biol. 24:2274-80 (2014)).
Glutaminases are encoded by two different genes: GLS1, which encodes kidney-type glutaminase (GLS), and GLS2, which encodes liver-type glutaminase. There are also two isoforms of GLS: one that is commonly referred to as kidney-type glutaminase (KGA) and the other, a C-terminal splice variant, designated glutaminase C (GAC). Because GAC is often highly expressed in cancer cells, it has been described as a “gate-keeper” enzyme for the elevated glutamine metabolism exhibited by these cells (“glutamine addiction”). Thus, GAC, as well as the longer GLS isoform KGA, are attractive therapeutic targets (Lukey et al., “Therapeutic Strategies Impacting Cancer Cell Glutamine Metabolism,” Future Med. Chem. 5:1685-700 (2013); van den Heuvel et al., “Analysis of Glutamine Dependency in Non-Small Cell Lung Cancer,” Cancer Biol. Ther. 13:1185-94 (2012)).
Currently, there are three classes of inhibitors for KGA and GAC. One class is represented by the benzophenanthridines, and, specifically, compound 968, which was previously demonstrated to act as a non-competitive allosteric inhibitor of GAC by interfering with its ability to undergo normal monomer-monomer interactions that lead to GAC dimers and, ultimately, to activated tetramers (Wang et al., “Targeting Mitochondrial Glutaminase Activity Inhibits Oncogenic Transformation,” Cancer Cell 18:207-19 (2010); Stalnecker et al., “Mechanism by Which a Recently Discovered Allosteric Inhibitor Blocks Glutamine Metabolism in Transformed Cells,” Proc. Natl. Acad. Sci. U.S.A. 112:394-99 (2015); Katt et al., “Dibenzophenanthridines as Inhibitors of Glutaminase C and Cancer Cell Proliferation,” Mol. Cancer Ther. 11:1269-78 (2012)). A second class of KGA/GAC inhibitors consists of analogs of the substrate glutamine, such as diazo-O-norleucine (DON), which binds to the enzyme active site and covalently modifies the catalytic serine (Ser-291) (Thangavelu et al., “Structural Basis for the Active Site Inhibition Mechanism of Human Kidney-Type Glutaminase (KGA),” Sci. Rep. 4:3827 (2014)). The third class of inhibitors, depicted in FIG. 1, consists of a number of bisthiadiazole derivatives, the prototype being bis-2-(5-phenylacetamido-1,2,4-thiadiazol-2-yl)ethyl (BPTES) (Thangavelu et al., “Structural Basis for the Allosteric Inhibitory Mechanism of Human Kidney-Type Glutaminase (KGA) and Its Regulation by Raf-Mek-Erk Signaling in Cancer Cell Metabolism,” Proc. Natl. Acad. Sci. U.S.A. 109:7705-10 (2012); Robinson et al., “Novel Mechanism of Inhibition of Rat Kidney-Type Glutaminase by Bis-2-(5-Phenylacetamido-1,2,4-Thiadiazol-2-yl)Ethyl Sulfide (BPTES),” Biochem. J. 406:407-14 (2007); Hartwick & Curthoys, “BPTES Inhibition of hGA124-551, a Truncated Form of Human Kidney-Type Glutaminase,” J. Enzyme Inhib. Med. Chem. 27:861-67 (2012); DeLaBarre et al., “Full-Length Human Glutaminase in Complex with an Allosteric Inhibitor,” Biochemistry 50:10764-70 (2011); Cassago et al., “Mitochondrial Localization and Structure-Based Phosphate Activation Mechanism of Glutaminase C with Implications for Cancer Metabolism,” Proc. Natl. Acad. Sci. U.S.A. 109:1092-97 (2012)). Gross et al. (Gross et al., “Antitumor Activity of the Glutaminase Inhibitor CB-839 in Triple-Negative Breast Cancer,” Mol. Cancer Ther. 13:890-901 (2014)) have described a BPTES derivative, CB-839, which is a more potent inhibitor than BPTES, and showed it to be effective against triple-negative breast cancer cells. CB-839 efficacy has been examined in vivo and subsequently tested in clinical trials (Matre et al., “Efficacy of Novel Glutaminase Inhibitor CB-839 in Acute Myeloid Leukemia,” Blood 124:3763 (2014)).
The discovery of BPTES as an inhibitor of KGA/GAC activity was first reported by Robinson et al. (Robinson et al., “Novel Mechanism of Inhibition of Rat Kidney-Type Glutaminase by Bis-2-(5-Phenylacetamido-1,2,4-Thiadiazol-2-yl)Ethyl Sulfide (BPTES),” Biochem. J. 406:407-14 (2007)). Elucidation of the binding site of BPTES, based on the X-ray crystal structure solved for the inhibitor bound to the KGA/GAC enzymes, revealed that its interactions with a flexible loop within the dimer-dimer interface of the tetrameric forms of these enzymes (i.e., the “activation loop”), accounted for its mode of inhibition. Indeed, mutations along this loop (316KEPSGLRFNKLF327 (SEQ ID NO:1); unless expressly stated otherwise, the amino acid numbering used in this application is in reference to mouse GAC) can markedly impact enzyme activity. The functional consequences of these mutations vary from inducing constitutive activation in the absence of phosphate (K325A), to shifting the dose response for phosphate (F322Y/F327S, K316A), to inhibiting the formation of higher-order oligomers (K316Q) (DeLaBarre et al., “Full-Length Human Glutaminase in Complex with an Allosteric Inhibitor,” Biochemistry 50:10764-70 (2011); McDonald et al., “Effect of Lysine to Alanine Mutations on the Phosphate Activation and BPTES Inhibition of Glutaminase,” Neurochem. Int 88:10-14 (2015); Ferreira et al., “Active Glutaminase C Self-Assembles into a Supratetrameric Oligomer That Can Be Disrupted by an Allosteric Inhibitor,” J. Biol. Chem. 288:28009-20 (2013)). Therefore, probing the conformation of this loop and how small molecules affect its orientation should provide a more detailed understanding of the fundamental mechanisms underlying the activation of the GLS isoforms.
The technology described herein is directed to overcoming these and other deficiencies in the art.